INTRODUCTION
Beginning in the 1970s, tremendous strides were made by the St. Lucian government in collaboration with the Rockefeller Foundation in the control of human intestinal schistosomiasis caused by Schistosoma mansoni, and by 1981, the prevalence had been reduced from 17% to < 2%.1 This resulted in control activities being sharply reduced without elimination of the disease.2 Currently, the transmission of S. mansoni is still considered ongoing in St. Lucia,3–5 although a recent study by Gaspard et al.6 suggests that transmission may have been interrupted after finding no positive cases in schoolchildren living adjacent to potential transmission sites. The intermediate host snail, Biomphalaria glabrata, is assumed to be present because efforts to eliminate the snail using Melanoides tuberculata, an exotic snail species as a biological competitor agent, may not have been completely successful.7 Another potential intermediate host snail, Biomphalaria straminea, was reported for the first time on the island in 1992, although its role in the transmission of schistosomiasis in St. Lucia is unknown.7 Other Biomphalaria species such as B. kuhniana, known from other Caribbean islands, might also be present.8 Gaspard et al.6 have rightly pointed out that “the situation in St. Lucia presents an opportunity to develop and evaluate possible approaches for verifying interruption of transmission.” A first step is to assess the current state of S. mansoni infection on Saint Lucia, and this should include both in humans and the snail intermediate host, B. glabrata (and possibly other Biomphalaria species present).
Thus, the government of St. Lucia established a steering committee with representatives from national key stakeholders under the coordination of the Ministry of Health and Wellness with the assistance of the Pan American Health Organization to establish guidelines to confirm interruption of transmission based on available information.6 Hence, updated information on the occurrence and geographic distribution of the intermediate hosts of S. mansoni (Biomphalaria spp.) and their infection status, based on a national malacological survey, is necessary to inform the guidelines. This information combined with national schistosomiasis surveys focusing on schoolchildren and other risk groups will give a more complete picture of the island’s current schistosomiasis status, needed to form the basis for an informed decision going forward. The inclusion of surveillance of known or suspected intermediate hosts, and their infection status in schistosomiasis control programs has become more pertinent, especially in areas where near elimination status of schistosomiasis needs to be confirmed.9
To inform the St. Lucian government further with respect to the status of schistosomiasis on the island, we conducted a targeted national malacological survey in St. Lucia in June–July 2022 to determine the geographic distribution and infection status of intermediate host snail(s) of S. mansoni, and to ascertain whether transmission might be ongoing. Here, we present the results of our study including updated geographic distribution data and trematode infection status for relevant snail species, employing both classical and molecular-based identification criteria. The malacological survey was undertaken in parallel with an environmental DNA (eDNA)-based sampling program designed to detect S. mansoni eDNA in snail habitats, a method shown to be highly sensitive for the detection of S. mansoni even in low-transmission settings.10 Additionally, we also undertook experimental infections of Biomphalaria species recovered from St. Lucia with S. mansoni to assess their status as potential mediators of transmission. Results from this study will help form the basis for recommendations for future efforts to monitor the interruption of schistosomiasis transmission, potentially including declarations of elimination of S. mansoni in St. Lucia.
MATERIALS AND METHODS
Collection of snails and screening for trematode cercariae.
We collected snails from 58 freshwater habitats on the island of St. Lucia between 25 June and 8 July 2022 (Figure 1 and Supplemental Table 1). Habitats that are overlapping in Figure 1 can be identified more fully in Supplemental File 1. At each habitat we recorded GPS coordinates, habitat type, distinctive features, the presence of people and/or animals, collected water samples appropriate for eDNA analysis (discussed subsequently), and then collected freshwater snails. Aquatic snails were collected along the water’s edge using kitchen sieves to sweep aquatic vegetation or a long-handled metal net to scoop along the substrate, rocks, and aquatic vegetation in deeper water. Snails were also picked off submerged rocks, plants, sticks, or debris using forceps. Collection at each habitat lasted between 30 minutes to 1 hour. Additional time was spent in the single habitat positive for Biomphalaria glabrata.
ExpertGPS basemap of St. Lucia and the habitats we sampled for aquatic snails. On the left side of the pie charts yellow indicates habitats that harbored B. kuhniana, red B. glabrata, and black no Biomphalaria. On the right side of the pie charts blue indicates eDNA samples were taken and white no samples were taken. See key on map for color coded pie charts.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
ExpertGPS basemap of St. Lucia and the habitats we sampled for aquatic snails. On the left side of the pie charts yellow indicates habitats that harbored B. kuhniana, red B. glabrata, and black no Biomphalaria. On the right side of the pie charts blue indicates eDNA samples were taken and white no samples were taken. See key on map for color coded pie charts.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
ExpertGPS basemap of St. Lucia and the habitats we sampled for aquatic snails. On the left side of the pie charts yellow indicates habitats that harbored B. kuhniana, red B. glabrata, and black no Biomphalaria. On the right side of the pie charts blue indicates eDNA samples were taken and white no samples were taken. See key on map for color coded pie charts.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
After collection, snails were cleaned with a Kimwipe™ (Kimberly-Clark Global Sales, LLC, Roswell, GA) to remove debris on their shells and then rinsed with clean water. Snails were placed individually into 12-well tissue culture plates in 3 mL of rainwater. Tissue culture plates were placed in ambient light for 1 hour to induce shedding of cercariae and were then screened for cercariae using an Olympus SZ61 (Tokyo, Japan) dissecting microscope. Keys based on shell or anatomic characters were used for the identification of snails11 and the cercariae they released.12,13 Snails and cercariae were preserved in 95% ethanol for later molecular analysis. Some snails were relaxed using menthol crystals, following the procedures of Pan,14 and were removed from their shells and fixed in Raillet–Henry’s solution to facilitate dissections and anatomical observations.
Molecular characterization of Biomphalaria and trematodes.
One to three snails from habitats that harbored Biomphalaria were used for DNA extraction (SL1–SL15) (Table 1). A portion of the ovotestis cut from the whole snail using a razor blade was used for DNA extraction, and the rest of the snail was put in 95% ethanol and vouchered in the Museum of Southwestern Biology Parasite Division (MSB:HOST). Snail genomic DNA was extracted using the ENZA Mollusc Kit (Omega Bio-Tek, Norcross, GA). The elution volume was 60 μL, and the elution buffer was allowed to soak on the filter for 5 minutes before centrifugation. The 16S rRNA gene was chosen for amplification and sequencing due to the diversity of specimens in GenBank for Neotropical Biomphalaria for phylogenetic and genetic distance comparisons. Partial sequences of the 16S rRNA gene were amplified by polymerase chain reaction (PCR) using the primers published by Palumbi15: 16Sar, 5′-CGCCTGTTTATCAAAAACAT-3′ and 16Sbr, 5′-CCGGTCTGAACTCAGATCACGT-3′. The volume of each PCR was 25 μL, with 100 ng of DNA, 0.8 mM/L of deoxynucleotides, 2.5 mM/L of MgCl2, 0.25 units of TaKaRa Ex Taq DNA polymerase (Clontech, Mountain View, CA), and 0.15 μM/L of each primer. PCR cycles followed the study of Palumbi,15 with the exception of the annealing temperature of 44.1°C and run on an Eppendorf Mastercycler ep gradient S.
Biomphalaria specimens and trematodes were chosen for DNA extraction and sequencing to determine species. Habitat, museum voucher numbers, and GenBank accession numbers are listed for each specimen
| Sample | Species | Habitat | Musuem voucher | GenBank accession number(s) |
|---|---|---|---|---|
| SL1 | B. kuhniana | Jacmel Gutter | MSB:Host:24883 | OQ862773 |
| SL2 | B. kuhniana | Jacmel Gutter | MSB:Host:24878 | OQ862774 |
| SL3 | B. kuhniana | Garrand-Babonneau | MSB:Host:24884 | OQ862275 |
| SL4 | B. kuhniana | Anse La Raye Primary School Gutter | MSB:Host:24885 | OQ862776 |
| SL5 | B. kuhniana | Anse La Raye Primary School Gutter | MSB:Host:24879 | OQ862777 |
| SL6 | B. kuhniana | Union Ministry Health Ditch | MSB:Host:24887 | OQ862778 |
| SL7 | B. kuhniana (juvenile) | Belmont of Grande Riviere | MSB:Host:24876 | OQ862779 |
| SL8 | B. kuhniana (juvenile) | Belmont of Grande Riviere | MSB:Host:24880 | OQ862780 |
| SL9 | B. kuhniana | Latille Falls Micoud | MSB:Host:24888 | OQ862781 |
| SL10 | B. kuhniana | Beausejour | MSB:Host:24881 | OQ862782 |
| SL11 | B. kuhniana | Beausejour | MSB:Host:24874 | OQ862783 |
| SL12 | B. glabrata | Cul de Sac Marsh | MSB:Host:24875 | OQ862784 |
| SL13 | B. glabrata | Cul de Sac Marsh | MSB:Host:24886 | OQ862785 |
| SL14 | B. kuhniana | Laborie | MSB:Host:24882 | OQ862786 |
| SL15 | B. kuhniana (Adult) | Belmont of Grande Riviere | MSB:Host:24877 | OQ862787 |
| SL16 | Schistosomatidae E | Belmont of Grande Riviere | MSB:Para:35977 | OQ868122; OQ866318 |
| SL17 | Patagifer sp. 2 | Belmont of Grande Riviere | MSB:Para:35976 | OQ868123; OQ868519 |
| SL18 | Patagifer sp. 2 | Cul de Sac Marsh | MSB:Para:35978 | OQ868124; OQ868518 |
Genomic DNA from one or two trematode cercariae collected from shedding snails (Table 1) were extracted using the Qiagen DNA Micro Kit (Qiagen, Valencia, CA) with a final elution volume of 35 μL. The 28S gene was amplified using forward primer, dig12 (5′-AAGCATATCACTAAGCGG-3′) and reverse primer 1500R (5′-GCTATCCTGAGGGAAACTTCG-3′).16 The volume of each PCR reaction was 25 μL with 2 μL of 50 ng of DNA, 0.8 mM/L dNTPs,2.5 mM/L MgCl2, 0.25 units of Ex Taq DNA (Clontech, Mountain View, CA), and 0.4 μM/L of each. The cox1 gene for the avian schistosome was amplified using the Schist 5′ and Schist 3′ primers.17 The nad1 gene for the echinostomes were amplified using the forward primer NDJ11 (5′-AGA TTCGTA AGG GGC CTA ATA-3′) and the reverse primer NDJ2a (5′-CTT CAG CCT CAG CAT AAT-3′)18 with the same PCR setup as the 28S gene. See Laidemitt et al.19 for PCR profiles of the nad1 and 28S genes.
PCR products were separated by agarose gel electrophoresis, visualized with a 0.5% GelRed nucleic acid gel stain (Biotium Inc., Hayward, CA) and were purified using ExoSap-IT (Applied Biosystems, Foster City, CA). Both strands were sequenced using an Applied Biosystems 3130 automated sequencer and BigDye Terminator Cycle Sequencing Kit version 3.1 (Applied Biosystems). DNA sequences were verified by aligning reads from the 5′ and 3′ directions using Sequencher 5.1 and manually corrected for ambiguous base calls (Gene Codes, Ann Arbor, MI). Approximately 460 bases were generated of the 16S rRNA gene. Sequences were aligned by CLUSTAL W, and the best fit model of substitution was modeled in Molecular Evolutionary Genetics Analysis 11 (MEGA11).20 Phylogenetic analyses using maximum likelihood (ML) included our 15 samples along with 56 sequences from the National Center for Biotechnology Information-GenBank for 16S. A total of 460 positions were used and along with Heuristic searches, 1,000 bootstrap replicates were run in MEGA11 for ML analysis. Uncorrected pairwise distance values (P values) were calculated in MEGA11, and a > 5% in mtDNA difference between samples was used to provisionally delineate species.21,22
Detecting S. mansoni in snails using a nad5 PCR assay.
A nicotinamide adenine dinucleotide dehydrogenase subunit 5 (nad5) PCR assay23 was used on the extracted DNA Biomphalaria samples (SL1–SL15) to determine if there were prepatent S. mansoni infections that could be detected by PCR. This is a sensitive assay (> 0.1 fg DNA) and differentiates Schistosoma species either by band size or absence/presence. We followed the same PCR and gel imaging protocol as Lu et al.23 except we used TaKaRa Ex Taq DNA® polymerase, buffer, and dNTPs (Clontech, Mountain View, CA). For a positive control, we used DNA from one Biomphalaria glabrata shedding S. mansoni.
Sampling and analysis of water to detect S. mansoni eDNA.
Water was collected from the snail habitats by a person designated to collect water samples for eDNA analysis, before anyone disturbing the habitat. This was done to avoid any potential contamination with parasite DNA of the water sample. A representation of the collected samples (Figure 1) covering the habitats where Biomphalaria snails were collected as well as geographically spread out on St. Lucia were selected for further eDNA analysis. Three replicates of 500 mL of water were collected from each habitat, with several distinct areas (both sides of a stream, vegetated and unvegetated, etc.) per habitat depending on the total area of each habitat. Bottles containing water samples were labeled appropriately and kept cool until returned to the laboratory, where the water sample was filtered the same day as collection. To capture eDNA, each sample was filtered using a vacuum pump (Chemical, Duty Pump, Millipore, Billerica, MA) through a 0.45-µm disc-filter (Whatman® glass microfiber filters WHA1825047, or Whatman (Sigma-Aldrich, Milwaukee, WI) membrane filters 7140-104). If the water was extremely turbid, a larger pore filter of 2.7 µm was used (Whatman glass microfiber filters, WHA1823047). The filter was then removed and placed in a separate LoBind 2-mL Eppendorf tube with RNAlater and held at room temperature until the filter was prepared for eDNA analysis. The entire filtering apparatus and collection bottles were cleaned with 5% bleach and rinsed three times between samples to avoid cross contamination.
In brief, DNA was extracted from the filters using the DNeasy Blood and Tissue Kit (Qiagen) following a modified protocol for water eDNA samples, as described in Spens et al.24 Extraction blanks were included for all extractions. A qPCR-assay with species-specific primers/probe targeting S. mansoni was used for analyzing the samples in PCR triplicates, following Sengupta et al.,10 where details on PCR mastermix, and thermal settings are found. Negative controls (NTC) were included for all qPCR runs. Filters with S. mansoni cercariae ranging from 1 to 25 cercariae were used as positive controls and DNA was extracted from the filters and analyzed with the S. mansoni specific quantitative PCR assay in the same way as the water samples. Lastly, to check for inhibition in the water samples, an internal positive control (TaqMan Exogeneous Internal Positive Control) was included for all samples.
Experimental exposures of field-derived and laboratory-reared St. Lucian Biomphalaria to S. mansoni.
Biomphalaria glabrata from Cul de Sac and B. kuhniana from Anse la Raye Primary School, Beausejour, and Belmont Grande of Riviere collecting habitats were used to establish laboratory colonies at the University of New Mexico (UNM). Both field-caught and F1 laboratory-reared snails (all initially negative for schistosome or other trematode infections as ascertained by lack of shedding cercariae) were exposed to miracidia of the Puerto Rico 1 (PR1) strain of S. mansoni originally sourced from the Schistosomiasis Resource Center, Biomedical Research Institute, Rockville, MD. Additionally, F1 generation snails were exposed to miracidia from a Kenyan isolate of S. mansoni maintained since 2013 at UNM in Biomphalaria choanomphala and hamsters. All snails were individually exposed to 10 freshly pipetted (hatched within 30 min) miracidia derived from livers of experimentally infected mice or hamsters and were maintained at 24°C, 12 h light:12 h dark cycle, fed red leaf lettuce ad libitum and shrimp pellets twice a week. They were examined for evidence of release of S. mansoni cercariae by isolating each snail in a well of a 12-well cell culture plate with 3 mL of artificial spring water and placing them under indirect light for 2 hours at 4.5, 5.5, 6.5, and 7.5 weeks post-exposure (WPE). All exposed snails were dissected after 7.5 WPE. Vertebrate animal use for this study was approved by the UNM Institutional Animal Care and Use Committee (IACUC 22-201290-MC).
In addition, some snails exposed to S. mansoni for 1, 2, or 4 days were prepared for histological examination. Snails were placed in Railliet–Henry’s fixative for at least 48 hours. The shell of each snail was removed, and the head-foot of the snail was placed in 10% neutral buffered formalin. The snails were processed at TriCore Reference Laboratories in Albuquerque, New Mexico, sectioned, and sections stained with hematoxylin and eosin.
RESULTS
Snail and trematode molecular results.
Among the 58 habitats sampled (Figure 1), we collected freshwater snails representing at least 11 species, all of which were isolated and examined for trematode cercariae. All snail species and cercariae are reported in Supplemental Table 1. We recorded 659 Biomphalaria specimens from 11 habitats. Identification of the Biomphalaria snails was confirmed by examination of conchological features, dissections, and by sequence data for the 16S rRNA marker gene. Biomphalaria glabrata was recovered from a single habitat (Cul de Sac) (Figure 2). All other Biomphalaria collected had shell anatomy and size, dissected genitalia including numbers of prostate diverticuli, and sequence data consistent with B. kuhniana as described by Pointier.11
Evolutionary analysis by maximum likelihood (ML). The ML tree was based on 460 positions of the 16S rRNA gene from 15 specimens of Biomphalaria collected in this study (bolded) and 56 specimens from GenBank. The B. glabrata specimens are in red and the B. kuhniana specimens are in green. Sister species in the genera Planorbella were chosen for the outgroup. A total of 1,000 bootstraps were run, and model GTR + I + G was selected via model selection. The percentage of trees in which the associated taxa clustered together is shown next to the branches (> 90%). Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood approach, and then selecting the topology with superior log likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Codon positions included were 1st+2nd+3rd+Noncoding. Evolutionary analyses were conducted in MEGA11.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
Evolutionary analysis by maximum likelihood (ML). The ML tree was based on 460 positions of the 16S rRNA gene from 15 specimens of Biomphalaria collected in this study (bolded) and 56 specimens from GenBank. The B. glabrata specimens are in red and the B. kuhniana specimens are in green. Sister species in the genera Planorbella were chosen for the outgroup. A total of 1,000 bootstraps were run, and model GTR + I + G was selected via model selection. The percentage of trees in which the associated taxa clustered together is shown next to the branches (> 90%). Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood approach, and then selecting the topology with superior log likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Codon positions included were 1st+2nd+3rd+Noncoding. Evolutionary analyses were conducted in MEGA11.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
Evolutionary analysis by maximum likelihood (ML). The ML tree was based on 460 positions of the 16S rRNA gene from 15 specimens of Biomphalaria collected in this study (bolded) and 56 specimens from GenBank. The B. glabrata specimens are in red and the B. kuhniana specimens are in green. Sister species in the genera Planorbella were chosen for the outgroup. A total of 1,000 bootstraps were run, and model GTR + I + G was selected via model selection. The percentage of trees in which the associated taxa clustered together is shown next to the branches (> 90%). Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood approach, and then selecting the topology with superior log likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Codon positions included were 1st+2nd+3rd+Noncoding. Evolutionary analyses were conducted in MEGA11.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
None of the Biomphalaria snails collected from St. Lucia in this study were infected with S. mansoni. One B. kuhniana from Belmont of Grande Ravine in the Parish of Dennery was shedding schistosome cercariae, which proved to have eyespots and to be cercariae of an avian-infecting schistosome species. Examination of 28S and cox1 gene markers indicated these St. Lucian avian schistosome cercariae were very similar to cercariae from B. straminea from Brazil designated as Schistosomatidae E.25 These cercariae are likely congeners to an avian schistosome (AY829246) from Biomphalaria sudanica from Kenya.26 One B. kuhniana from Belmont of Grande Ravine and three B. glabrata from Cul de Sac were shedding Patagifer sp. 2 (based on 28S and nad1 gene markers). This species is known to infect two Biomphalaria species in Kenya,19 and a similar cercaria has been reported from Biomphalaria orbignyi from Argentina.27 The genus is known for infecting ibises (subfamily Threskiornithinae) as definitive hosts and snails as first and second intermediate hosts.28 No other trematode infections were found among the St. Lucian Biomphalaria snails we sampled.
One of the most striking aspects of the freshwater snail fauna of St. Lucia is the extent to which it is dominated by the exotic snail, Melanodies tuberculata, which was collected from 49 of 58 habitats we sampled. In most places, particularly so for streams, it was by far the most abundant gastropod collected. Historically, before the introduction of M. tuberculata, B. glabrata was commonly recovered from streams. In the one marshy habitat where we found B. glabrata, M. tuberculata was also present but was not numerically dominant.
nad5 PCR assay.
No schistosome bands were detected in any of the 30 (four B. glabrata and 26 B. kuhniana) snails examined using this assay. Positive controls were positive, and negative controls were negative.
Schistosoma mansoni eDNA results.
A total of 63 filtered water samples from 21 locations (Table 1) were analyzed for S. mansoni eDNA, and no amplification of S. mansoni DNA was detected, including the Cul de Sac habitat where B. glabrata snails were found. All negative controls throughout the analysis process, such as extraction blanks and PCR negative controls, for example, were negative, and all the positive controls were also positive. For the filters spiked with S. mansoni cercariae, the qPCR assay picked up an S. mansoni DNA signal on each filter, even the filter containing only one S. mansoni cercariae. Amplification of the internal positive control indicated that no inhibition was detected in the water samples.
Experimental exposures.
With respect to experimental exposures to S. mansoni (Table 2), B. glabrata from Cul de Sac proved to be susceptible to both PR1 (Puerto Rican) and Kenyan isolates of S. mansoni, particularly so to the isolate of Caribbean origin, which produced higher infection rates (61%) than the Kenyan S. mansoni isolate (31%) at 7.5 WPE. The B. glabrata exposed to the PR1 isolate also started shedding cercariae a week earlier (4.5 WPE) compared with the B. glabrata exposed to the Kenyan isolate (5.5 WPE). None of the exposed B. glabrata that failed to shed cercariae of S. mansoni were found positive upon dissection. Also, the number of cercariae shed per infected snail, although not enumerated, were conspicuously fewer in snails exposed to S. mansoni of Kenyan origin. None of the B. kuhniana (field-derived or laboratory-reared F1s including neonates) exposed to either isolate of S. mansoni shed any S. mansoni cercariae. Examination of histological sections showed that S. mansoni miracidia penetrated B. kuhniana (Figure 3A), but the only sporocysts seen by 4 days postexposure were encapsulated and had lost their typical anatomical structure (Figure 3B). All B. kuhniana exposed to S. mansoni were dissected 48 days postexposure and were negative for sporocysts of S. mansoni.
Experimental exposure of Biomphalaria spp. with two strains of Schistosoma mansoni
| Species | Habitat | Origin of S. mansoni | No. of miracidia | No. of snails exposed | 4.5 WPE # infected/ survivors (% shedding) | 5.5 WPE # infected/ survivors (% shedding) | 6.5 WPE # infected/ survivors (% shedding) | 7.5 WPE # infected/ survivors (% shedding) | Dissections |
|---|---|---|---|---|---|---|---|---|---|
| B. glabrata (field-dervied 10–12 mm) | Cul de Sac Marsh | Puerto Rico | 10 | 22 | 2/20 (10%) | 8/20 (40%) | 9/20 (45%) | 11/18 (61%) 2/18 shed Patagifer | Snails dissected—11 positive and shedding cercariae |
| B. glabrata (F1s 5–7 mm) | Cul de Sac Marsh | Kenya | 10 | 22 | 0/18 (0%) | 2/16 (12.5%) | 4/16 (25%) | 5/16 (31%) | Snails dissected—only five positive and shedding cercariae |
| B. kuhniana (field-derived 6-8 mm) | Belmont of Grande Riviere | Puerto Rico | 10 | 24 | 0/23 (0%) | 0/23 (0%) | 0/23 (0%) | 0/18 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (neonates) | Belmont of Grande Riviere | Puerto Rico | 5 | 24 | 0/22 (0%) | 0/20 (0%) | 0/14 (0%) | 0/4 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (field-derived 6–8 mm) | Anse La Raye | Puerto Rico | 10 | 17 | 0/17 (0%) | 0/17 (0%) | 0/17 (0%) | 0/15 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (F1s 2–4 mm) | Anse La Raye | Kenya | 10 | 58 | 0/33 (0%) | 0/32 (0%) | 0/31 (0%) | 0/31 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (F1s 2–4 mm) | Anse La Raye | Puerto Rico | 10 | 48 | 0/36 (0%) | 0/36 (0%) | 0/36 (0%) | 0/36 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (Field-Derived 6–8 mm) | Stadium | Puerto Rico | 10 | 24 | 0/22 (0%) | 0/20 (0%) | 0/20 (0%) | 0/16 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (F1s 2–4 mm) | Beausejour | Kenya | 10 | 52 | 0/41 (0%) | 0/36 (0%) | 0/36 (0%) | 0/35 (0%) | Snails dissected negative for sporocysts or cercariae |
| B. kuhniana (F1s 2–4 mm) | Beausejour | Puerto Rico | 10 | 12 | 0/12 (0%) | 0/12 (0%) | 0/11 (0%) | 0/11 (0%) | Snails dissected negative for sporocysts or cercariae |
Histological sections of laboratory-reared Biomphalaria kuhniana from Anse La Raye, St. Lucia, exposed to 10 PR1 S. mansoni miracidia for 1 day post-exposure (DPE) (A) or 4 DPE (B). At 1 DPE, the sporocyst retains its anatomic integrity and a thin layer of hemocytes can be seen surrounding it. At 4 DPE, the sporocyst has been heavily encapsulated by host hemocytes and its anatomical integrity lost.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
Histological sections of laboratory-reared Biomphalaria kuhniana from Anse La Raye, St. Lucia, exposed to 10 PR1 S. mansoni miracidia for 1 day post-exposure (DPE) (A) or 4 DPE (B). At 1 DPE, the sporocyst retains its anatomic integrity and a thin layer of hemocytes can be seen surrounding it. At 4 DPE, the sporocyst has been heavily encapsulated by host hemocytes and its anatomical integrity lost.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
Histological sections of laboratory-reared Biomphalaria kuhniana from Anse La Raye, St. Lucia, exposed to 10 PR1 S. mansoni miracidia for 1 day post-exposure (DPE) (A) or 4 DPE (B). At 1 DPE, the sporocyst retains its anatomic integrity and a thin layer of hemocytes can be seen surrounding it. At 4 DPE, the sporocyst has been heavily encapsulated by host hemocytes and its anatomical integrity lost.
Citation: The American Journal of Tropical Medicine and Hygiene 109, 4; 10.4269/ajtmh.23-0235
DISCUSSION
Biomphalaria glabrata is now rare, but not absent, on St. Lucia. It was found in only one of the 58 habitats we sampled. Of the 26 habitats sampled by Pointier 1993,7 our study sampled 21 of them. Of the remaining five, although we could not pinpoint the original locations specified by Pointier, we sampled habitats that were spatially close to them (Supplemental Table 1). Of the 21 habitats, 14 formerly harbored B. glabrata. Compared with precontrol efforts when B. glabrata was widely distributed across the island, its present distribution as best we could determine is limited to a single, but fairly extensive, flat marshland in northcentral St. Lucia. This Cul de Sac habitat was problematic for schistosomiasis transmission in St. Lucia in the past and was the last habitat known to be treated by molluscicide, which occurred in 1988.7
None of the Biomphalaria snails we collected on St. Lucia were infected with S. mansoni by shedding or PCR. However, it is well known that the infection prevalence/shedding rate of schistosome intermediate hosts can be quite low (often less than 2%), compared with the infection rates sometimes observed in mammalian hosts.29 Therefore, absence of infected snails among our samples does not necessarily rule out transmission. For this reason, we also deployed eDNA, a method that since it was first developed for S. mansoni,10,30 has found use as a sensitive, noninvasive environmental monitoring tool in medical and veterinary parasitology.31 None of the eDNA samples collected and analyzed on St. Lucia were positive for S. mansoni, which further supports the absence of infected snails in the identified Biomphalaria snail habitats (Figure 1). Especially in a low-transmission setting, such as would be expected in St. Lucia, the fact that S. mansoni eDNA can persist for up to 8 days10 further supports that no recent shedding events have taken place in the examined snail habitats. The absence of eDNA signals, whether from miracidia or cercariae, also suggests the infection is not being maintained in reservoir hosts, although this is an area that might require further study.
Although we did not find any indication of ongoing S. mansoni transmission, our experimental infection results indicated B. glabrata of Cul de Sac origin is still compatible with S. mansoni, as tested with the PR1 isolate of S. mansoni derived from Puerto Rico, which has been maintained for decades in the laboratory. It is also susceptible, although less so, with an isolate of S. mansoni from Kenya. It might be from an immigrant of African or southwest Asian origin whereby S. mansoni could potentially be reintroduced into St. Lucia, so the susceptibility of local B. glabrata to a Kenyan isolate highlights a lingering possibility for such an event.
The snail taxon we identified as B. kuhniana was relatively widely distributed on St. Lucia. It is appropriately cautious to consider our identification of the St. Lucian B. kuhniana specimens as provisional because it is clear from both the literature,32,33 including a recent detailed study by de Araujo et al.,34 and from GenBank entries that B. kuhniana, B. straminea, and B. intermedia are closely related and may best be viewed as a complex of diverging species. Genetic distances between some isolates of provisional B. kuhniana and B. straminea fall in an ambiguous region of < 3% divergence. Specimens indistinguishable from B. kuhniana based on appearance and sequence of our chosen marker gene were also found on Antigua and Montserrat.35 Further collection of specimens from broader geographic areas and additional sequence data from more genes, including nuclear genes, will be required to resolve this matter fully.
We show here that provisional B. kuhniana is capable of naturally transmitting an avian schistosome on St. Lucia. We are not aware of any reports of B. kuhniana serving as a natural host for S. mansoni, and Paraense32,36 indicated this snail had proven refractory to S. mansoni infection. Pointier11 considered its status as a host for S. mansoni to be “unclear.” Our experimental infections using both PR1 and Kenyan isolates of S. mansoni indicated that miracidia were able to penetrate B. kuhniana, and evidence of encapsulation of sporocysts and dismemberment by hemocytes was observed. Upon dissection of exposed snails at 48 days postexposure, none of the B. kuhniana showed indication of S. mansoni sporocysts or cercariae within them. We conclude the status of provisional B. kuhniana as potential future hosts of S. mansoni in the Caribbean should not be overlooked, but based on results from this study, they appear to be incompatible with S. mansoni development. Biomphalaria kuhniana is also a potential competitor of B. glabrata responsible in part for the latter species decline in abundance in Martinique,36,37 and it is conceivable that in certain situations, if co-occurring with B. glabrata, it could act as a decoy host for miracidia of S. mansoni.
The current abundance of Melanoides tuberculata in almost all freshwater habitats on St. Lucia is remarkable, and it is certainly tempting to conclude that this species has played a major role in diminishing B. glabrata abundance on the island by playing a competitor role previously discussed by several authors.7,11,38,39 In a previous survey of 26 St. Lucian snail habitats, Pointier7 noted that 24 (92.3%) habitats contained M. tuberculata, and 19 (73.0%) harbored B. glabrata, with 17 (65.4%) containing both species. The two habitats where B. glabrata was most abundant lacked M. tuberculata. Comparison with our results suggests that B. glabrata has since become much less common on St. Lucia, and even the single habitat where we found B. glabrata also harbored M. tuberculata.
Although beneficial from the standpoint of schistosomiasis control in the short term, the presence of M. tuberculata is also reason for concern because it represents a major alteration to native St. Lucian freshwater biotas, similar to what has happened on Martinique.36 Such snails might begin to transmit trematodes of medical, veterinary, or conservation concern.40,41 Also, it should be noted that the flora and fauna of islands are subject to high immigration and extinction rates, so future changes in freshwater habitats in St. Lucia may be unpredictable, particularly considering changing climates.
In conclusion, our study provides no evidence for the continued presence of S. mansoni on St. Lucia. Both snail and eDNA surveys found no evidence for the presence of S. mansoni. Furthermore, compared with past surveys, we found that the primary snail vector of concern, B. glabrata, is now rare on St. Lucia, and that the other Biomphalaria species present, B. kuhniana, does not support the full intramolluscan development of S. mansoni culminating in production of human-infecting cercariae or patent infections. In fact, B. kuhniana seems to mount active resistance responses that destroy young S. mansoni sporocysts. We note that the susceptibility status for S. mansoni of the snails we provisionally identified as B. kuhniana could change, possibly influenced by its infection with other facilitating parasite species,42 or in response to relaxed selection for resistance owing to the present lack of S. mansoni. Additionally, while visiting freshwater habitats across the island, including Cul de Sac, we were struck by the limited extent to which St. Lucians now seem to depend on surface waters for bathing or clothes washing, thereby diminishing the likelihood of exposure to infection should the parasite be present. Finally, we note that the possibility remains that there are isolated foci of infection maintained by rodent or other reservoir hosts,43–45 though this seems less likely to be an issue of concern given the rarity of B. glabrata, the lack of supporting eDNA evidence, and the potential difficulty of short-lived rodents to maintain transmission under such circumstances.
Supplemental Materials
ACKNOWLEDGMENTS
Authors give special acknowledgment to staff from the Ministry of Health, Wellness and Environment and the Ministry of Agriculture, Fisheries, Forestry, Food Security and Rural Development of St. Lucia who assisted with the field work. The authors thank the Biomedical Research Institute for providing the PR1 S. mansoni eggs. The reagents were provided by the National Institute of Allergy and Infectious Diseases (NIAID) Schistosomiasis Resource Center of the Biomedical Research Institute (Rockville, MD) through NIH-NIAID Contract HHSN272201700014I. We also thank Ms. Karen Buehler from Tri-Core Laboratories in Albuquerque, NM, for her timely assistance in producing histological sections of snails and Dr. Sharmine Melville-Edwin (Chief Veterinary Officer), Ministry of Agriculture, Fisheries, Forestry, Food Security and Rural Development, Union, Castries, Saint Lucia for her assistance in organizing the logistics of the field work.
REFERENCES
- 1.↑
Jordan P , 1985. Schistosomiasis: The St. Lucia Project. Cambridge, United Kingdom: Cambridge University Press.
- 2.↑
Pan American Health Organization , 2014. Schistosomiasis Regional Meeting. Defining a Road Map toward Verification of Elimination of Schistosomiasis Transmission in Latin America and the Caribbean by 2020. Washington, DC: PAHO.
- 3.↑
Kurup R , Hunjan GS , 2010. Epidemiology and control of schistosomiasis and other intestinal parasitic infections among school children in three rural villages of south Saint Lucia. J Vector Borne Dis 47: 228–234.
- 4.↑
Hewitt R , Willingham AL , 2019. Status of schistosomiasis elimination in the Caribbean region. Trop Med Infect Dis 4: 24.
- 6.↑
Gaspard J , Usey MM , Fredericks-James M , Sanchez-Martin M , Atkins L , Campbell Jr CH , Corstjens PLAM , van Dam GJ , Colley DG , Secor WE , 2020. Survey of schistosomiasis in Saint Lucia: evidence for interruption of transmission. Am J Trop Med Hyg 102: 827–831.
- 7.↑
Pointier JP , 1993. The introduction of Melanoides tuberculata (Mollusca: Thiaridae) to the island of Saint Lucia (West Indies) and its role in the decline of Biomphalaria glabrata, the snail intermediate host of Schistosoma mansoni. Acta Trop 54: 13–18.
- 8.↑
Pointier JP , Samadi S , Jarne P , Delay B , 1998. Introduction and spread of Thiara granifera (Lamarck, 1822) in Martinique, French West Indies. Biodivers Conserv 7: 1277–1290.
- 9.↑
Rollinson D et al., 2013. Time to set the agenda for schistosomiasis elimination. Acta Trop 128: 423–440.
- 10.↑
Sengupta ME et al., 2019. Environmental DNA for improved detection and environmental surveillance of schistosomiasis. Proc Natl Acad Sci USA 116: 8931–8940.
- 11.↑
Pointier JP , 2008. Guide to the Freshwater Molluscs of the Lesser Antiles. Hackenheim, Germany: Conch Books.
- 12.↑
Frandsen F , Christensen NO , 1984. An introductory guide to the identification of cercariae from African freshwater snails with special reference to cercariae of trematode species of medical and veterinary importance. Acta Trop 41: 181–202.
- 13.↑
Schell SC , 1985. Handbook of Trematodes of North America North of Mexico. Boise, ID: University of Idaho Press.
- 14.↑
Pan CT , 1958. The general histology and topographic micranatomy of Australorbis glabratus. Bull Mus Comp Zool Harv 119: 235–299.
- 16.↑
Tkach VV , Littlewood DTJ , Olson PD , Kinsella JM , Swiderski Z , 2003. Molecular phylogenetic analysis of the Microphalloidea Ward, 1901 (Trematoda: Digenea). Syst Parasitol 56: 1–15.
- 17.↑
Lockyer AE et al., 2003. The phylogeny of the Schistosomatidae based on three genes with emphasis on the interrelationships of Schistosoma Weinland, 1858. Parasitology 126: 203–224.
- 18.↑
Kostadinova A , Herniou EA , Barrett J , Littlewood DT , 2003. Phylogenetic relationships of Echinostoma Rudolphi, 1809 (Digenea: Echinostomatidae) and related genera re-assessed via DNA and morphological analyses. Syst Parasitol 54: 159–176.
- 19.↑
Laidemitt MR , Brant SV , Mutuku MW , Mkoji GM , Loker ES , 2019. The diverse echinostomes from East Africa: with a focus on species that use Biomphalaria and Bulinus as intermediate hosts. Acta Trop 193: 38–49.
- 20.↑
Tamura K , Stecher G , Kumar S , 2021. MEGA11: molecular evolutionary genetics analysis version 11. Mol Biol Evol 38: 3022–3027.
- 21.↑
Vilas R , Criscione CD , Blouin MS , 2005. A comparison between mitochondrial DNA and the ribosomal internal transcribed regions in prospecting for cryptic species of platyhelminth parasites. Parasitology 131: 839–846.
- 22.↑
Detwiler JT , Bos DH , Minchella DJ , 2010. Revealing the secret lives of cryptic species: examining the phylogenetic relationships of echinostome parasites in North America. Mol Phylogenet Evol 55: 611–620.
- 23.↑
Lu L , Zhang SM , Mutuku MW , Mkoji GM , Loker ES , 2016. Relative compatibility of Schistosoma mansoni with Biomphalaria sudanica and B. pfeifferi from Kenya as assessed by PCR amplification of the S. mansoni ND5 gene in conjunction with traditional methods. Parasit Vectors 9: 166.
- 24.↑
Spens J , Evans AR , Halfmaerten D , Knudsen SW , Sengupta ME , Mak SST , Sigsgaard EE , Hellström M , 2017. Comparison of capture and storage methods for aqueous macrobial eDNA using an optimized extraction protocol: advantage of enclosed filter. Methods Ecol Evol 8: 635–645.
- 25.↑
Pinto HA , Mati VLT , Melo AL , Brant SV , 2022. A putative new genus of avian schistosome transmitted by Biomphalaria straminea (Gastropoda: Planorbidae) in Brazil, with a discussion on the potential involvement in human cercarial dermatitis. Parasitol Int 90: 102607.
- 26.↑
Brant SV , Bochte CA , Loker ES , 2011. New intermediate host records for the avian schistosomes Dendritobilharzia pulverulenta, Gigantobilharzia huronensis, and Trichobilharzia querquedulae from North America. J Parasitol 97: 946–949.
- 27.↑
Núñez M , Hamann M , Rumi A , 1997. Estudios de trematodes larvales en Biomphalaria spp. (Mollusca, Planorbidae) de la localidad de San Roque, provincia de Corrientes, Argentina. Physis 54: 7–15.
- 28.↑
Faltynkova A , Gibson DI , Kostadinova A , 2008. A revision of Petasiger Dietz, 1909 (Digenea: Echinostomatidae) and a key to its species. Syst Parasitol 71: 1–40.
- 29.↑
Nwoko OE , Kalinda C , Chimbari MJ , 2022. Systematic review and meta-analysis on the infection rates of schistosome transmitting snails in southern Africa. Trop Med Infect Dis 7: 72.
- 30.↑
Sato MO et al., 2018. Usefulness of environmental DNA for detecting Schistosoma mansoni occurrence sites in Madagascar. Int J Infect Dis 76: 130–136.
- 31.↑
Sengupta ME , Lynggaard C , Mukaratirwa S , Vennervald BJ , Stensgaard AS , 2022. Environmental DNA in human and veterinary parasitology—current applications and future prospects for monitoring and control. Food Waterborne Parasitol 29: e00183.
- 32.↑
Paraense W , 1988. Biomphalaria kuhniana (Clessin, 1883), planorbid mollusc from South America. Memorias Do Instituto Oswaldo Cruz. Mem Inst Oswaldo Cruz 83, 10.1590/S0074-02761988000100001.
- 33.↑
DeJong RJ et al., 2001. Evolutionary relationships and biogeography of Biomphalaria (Gastropoda: Planorbidae) with implications regarding its role as host of the human bloodfluke, Schistosoma mansoni. Mol Biol Evol 18: 2225–2239.
- 34.↑
de Araújo AD , Carvalho ODS , Gava SG , Caldeira RL , 2023. DNA barcoding as a valuable tool for delimiting mollusk species of the genus Biomphalaria Preston, 1910 (Gastropoda: Planorbidae). Front Cell Infect Microbiol 13: 1167787.
- 35.↑
Laidemitt MR , Buddenborg SK , Lewis LL , Michael LE , Sanchez MJ , Hewitt R , Loker ES , 2020. Schistosoma mansoni vector snails in Antigua and Montserrat, with snail-related considerations pertinent to a declaration of elimination of human schistosomiasis. Am J Trop Med Hyg 103: 2268–2277.
- 36.↑
Pointier JP , 2001. Invading freshwater snails and biological control in Martinique Island, French West Indies. Mem Inst Oswaldo Cruz 96: 67–74.
- 37.↑
Guyard A , Pointier JP , Théron A , Gilles A , 1982. Mollusques hôtes intermédiaires de la schistosomose intestinale dans les Petites Antilles. Hypothèses sur le rôle de Biomphalaria glabrata et B. straminea en Martinique. Malacologia 22: 103–107.
- 38.↑
Pointier JP , Facon B , Jarne P , David P , 2003. Thiarid snails, invading gastropods of tropical freshwater habitats. Xenophora 104: 14–20.
- 39.↑
Friani G , Ribeiro do Amaral AM , Quinelato S , Mello-Silva CC , Golo PS , 2023. Biological control of Biomphalaria, the intermediate host of Schistosoma spp.: a systematic review. Cienc Rural 53: e20210714.
- 40.↑
Lopes AS , Pulido-Murillo EA , Melo AL , Pinto HA , 2020. Haplorchis pumilio (Trematoda: Heterophyidae) as a new fish-borne zoonotic agent transmitted by Melanoides tuberculata (Mollusca: Thiaridae) in Brazil: a morphological and molecular study. Infect Genet Evol 85: 104495.
- 41.↑
Metz DCG , Turner AV , Nelson AP , Hechinger RF , 2023. Potential for emergence of foodborne trematodiases transmitted by an introduced snail (Melanoides tuberculata) in California and elsewhere in the United States. J Infect Dis 227: 183–192.
- 42.↑
Spatz L , Cappa SM , de Nunez MO , 2012. Susceptibility of wild populations of Biomphalaria spp. from neotropical South America to Schistosoma mansoni and interference of Zygocotyle lunata. J Parasitol 98: 1291–1295.
- 43.↑
Ketzis JK , Lejeune M , Branford I , Beierschmitt A , Willingham AL , 2020. Identification of Schistosoma mansoni infection in a nonhuman primate from St. Kitts more than 50 years after interruption of human transmission. Am J Trop Med Hyg 103: 2278–2281.
- 44.↑
Théron A , Pointier JP , Combes C , 1978. Approche écologique du problème de la responsibilité de l’homme et du rat dans le fonctionnement d’un site de transmission à Schistosoma mansoni en Guadeloupe. Ann Parasitol Hum Comp 53: 223–234.
- 45.↑
Theron A , 1984. Early and late shedding patterns of Schistosoma mansoni cercariae: ecological significance in transmission to human and murine hosts. J Parasitol 70: 652–655.